FEMS Microbiology Ecology 51 (2004) 19–29
www.fems-microbiology.org
Bacterial populations in the rhizosphere of tobacco plants
producing the quorum-sensing signals hexanoyl-homoserine
lactone and 3-oxo-hexanoyl-homoserine lactone
Cathy dÕAngelo-Picard a, Denis Faure a, Aurélien Carlier a, Stéphane Uroz a,
Aurélie Raffoux a, Rupert Fray b, Yves Dessaux a,*
a
Institut des Sciences du Végétal, Centre National de la Recherche Scientifique, UPR2355, Bâtiment 23, F-91198, Gif-sur-Yvette, France
b
School of BioSciences, University of Nottingham, Sutton Bonington Campus, Loughborough LE12 5RD, UK
Received 26 February 2004; received in revised form 21 June 2004; accepted 10 July 2004
First published online 20 August 2004
Abstract
A tobacco line genetically modified to produce two N-acyl homoserine lactones and its non-transformed parental line were grown
in non-sterile soil. Microbial populations inhabiting the bulk soil, and those colonizing the root system of the two tobacco lines,
were analyzed using cultivation-independent (phospholipid fatty acid and denaturing gradient gel electrophoresis) and cultivation-based assays. The cell density of total cultivable bacteria, fluorescent pseudomonads, sporulated, and thermotolerant bacteria
was also determined in a time-course experiment (15 weeks). A possible ‘‘rhizosphere effect’’ related to the development of the plant
was seen. However, no dissimilarities in cell population densities or population ratios of the microbial groups were detected in the
rhizosphere of the two plant lines. Similarly, bacterial communities that either produced N-acyl homoserine lactone or degraded the
signal hexanoyl homoserine lactone were enumerated from the two plant lines. No noticeable differences were evidenced from one
plant genotype to the other. Whilst the transgenic plants released detectable amounts of the quorum-sensing signal molecules and
efficiently cross-talked with the surrounding microbial populations, the bias generated by these signals in the reported experimental
conditions therefore appears to remain weak, if not non-existent.
2004 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved.
Keywords: Acyl homoserine lactone; Signal molecule; yenI; Root system; Transgenic plant; Engineered exudation
1. Introduction
Numerous bacteria, including some of those inhabiting the rhizosphere, have evolved a regulatory process
known as quorum sensing (QS) that allows them to
modulate gene expression synchronously and as a function of their cell density (for reviews, see [1,2]). Bacteria
that exhibit QS regulation produce limited amounts of
*
Corresponding author. Tel.: +33 1 6982 3690; fax: +33 1 6982
3695.
E-mail address: [email protected] (Y. Dessaux).
one or more signal molecules. The concentration of
the signal molecules increases as a function of cell density, up to a threshold concentration that permits its
sensing in the environment by the bacterial population.
The chemical nature of the QS signals used for cell-tocell communication varies according to the bacterial
species. In gram-negative bacteria, communication is
mostly mediated by N-acyl homoserine lactone molecules [1], abbreviated as N-AHSL.
QS-regulated functions in plant-associated microorganisms include pathogenicity, siderophore production,
plasmid conjugal transfer, production of antibiotics
0168-6496/$22.00 2004 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved.
doi:10.1016/j.femsec.2004.07.008
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C. dÕAngelo-Picard et al. / FEMS Microbiology Ecology 51 (2004) 19–29
and antifungal compounds, swarming, etc. [1]. Since
pathogenicity or pathogenicity-related functions are
controlled by QS in some major plant and animal pathogens (e.g. [3]), it has been proposed that the QS system could be an appropriate novel target against
which biological control agents [4] or drugs that attenuate and limit virulence [5,6] could be developed. Two
main strategies have been proposed. The first one,
which aimed at degrading the signal molecules soon
after their production, has been successfully demonstrated in many systems [5,7–9]. The second approach
proposes to saturate a given environment with NAHSL molecules [4,10,11], thus inducing bacteria to
generate an inappropriate sub-quorate response, detrimental to their own survival. Targeting the N-AHSL
synthase encoded by yenI (from Yersinia enterocolitica)
to the chloroplasts of transgenic tobacco plants resulted in the synthesis by plant cells of both hexanoyl
homoserine lactone (C6-HSL) and 3-oxo-hexanoyl
homoserine lactone (3-oxo-C6-HSL) signals. These
molecules generated a ‘‘cross-talk’’ amongst plants
and microorganisms, validating this approach
[4,10,11]. The production of N-AHSL by the plant
might, however, induce unexpected disturbances in
non-target microbial populations, including beneficial
ones. For example, the production of the N-AHSL
molecules by the plants [10] or by other bacteria [12] affected the biocontrol ability of Pseudomonas aureofaciens strains. It is therefore of prime importance to
evaluate what may be the consequence of the release
of such QS signals in the rhizosphere, on bacterial populations and communities.
The experiment reported here involved the transgenic
yenI tobacco line described above producing C6-HSL
and 3-oxo-C6-HSL. The density of cultivable bacteria
colonizing the plant root system was determined by
plating and counting, and assessed with reference to
the density of bacteria colonizing the root system of
wild-type (WT) plants. The analyzed bacteria belonged
to several microbial populations and communities were
chosen as indicator groups known to respond to
N-AHSL production (e.g. pseudomonads) or not or to
degrade the signal (e.g. sporulated). The density of bacteria implicated in QS regulation was also determined
by the direct assay of their ability to synthesize
N-AHSL or to quench the C6-HSL signal. Total populations and communities were also analyzed at the end
of the time course experiment with denaturing gradient
gel electrophoresis (DGGE) and phospholipid fatty acid
(PLFA) profiles of the root-associated microbial community. The experiment revealed that production of
QS molecules by the investigated transgenic tobacco line
neither drastically modifies the composition of the
microbial community, nor does it affect the incidence
of bacterial isolates implicated in QS regulation (producers or degraders).
2. Materials and methods
2.1. Plant and soil characteristics
Wild-type Nicotiana tabacum cv. Samson, and NAHSL-producing derivatives (T-plants) were cultured
under greenhouse confinement. The N-AHSL-producing tobacco harbors a T-DNA expressing the yenI gene
of Yersinia enterocolitica, the product of which directs
the synthesis of C6-HSL and 3-oxo-C6-HSL in a 1:1 ratio upon targeting to the chloroplasts [10]. Disinfected
seeds of WT and T-plant were germinated in a soil mixture (1/1; v/v) consisting of sterile Loire River sand and
unsterilized reference soil from La-Côte-Saint-André
(Isère, France) that was the source of rhizospheric bacteria. The reference soil was mostly loamy, with a pH
value of 5.8, a cation exchange capacity (CEC) of 108
cmol kg 1 and a nitrogen content of 0.28% [13]. The
pots (20 cm in diameter) were randomly placed in the
greenhouse, under long day conditions (16 h) at 17 C
(night) and 24 C (day), and watered daily with tap
water. Soil samples were collected at the beginning of
the experiment before planting (time zero, t0). Rhizosphere samples were collected after 7, 11 and 15 weeks.
At 7 weeks, plants were ca. 5 cm high, with ca. 6 leaves.
At 11 and 15 weeks, they were about 40/45 and 50/60 cm
high, and harbored 16/20 and 20/24 leaves, respectively.
At each sampling time (including time zero), five pots
(each with three tobacco plants except at time zero)
per condition were randomly chosen and the microbial
communities colonizing the soil or the mixed soil–root
systems of the three plants examined, except for PLFA
and DGGE analyses that were performed on three replicates (each with three plants) per condition.
All chemicals, including N-AHSL, were from commercial sources.
2.2. Enumeration of the cultivable rhizosphere microbial
community
From each of the pots, one gram of soil sample or
roots with adhering soil was used for determining the
number of cultivable cells. Soil moisture content was
determined by weighing fresh and dried soil (100 C
for 24 h). Soil or rhizospheric samples were resuspended
in 10 mL of sterile 0.8% NaCl by very vigourous shaking
for 3 min, and the resulting suspension was serially diluted. Appropriate dilutions were spread onto the following culture media containing 100 lg mL 1
cycloheximide: TY (0.5% tryptone, 0.3% yeast extract,
and 6 mM CaCl2) for total cultivable bacteria, spore
forming bacteria and thermotolerant bacteria, and
KBm modified from King et al. [14] supplemented with
ampicillin (40 lg mL 1) and chloramphenicol (13
lg mL 1) for fluorescent pseudomonads [15]. Plates
were incubated at 24 C in the dark, except for thermo-
C. dÕAngelo-Picard et al. / FEMS Microbiology Ecology 51 (2004) 19–29
21
tolerant bacteria that were incubated at 47 C. Sporulated bacteria were determined from dilutions performed from a boiled suspension (5 min at 100 C).
Colony forming units (CFU) were counted after 2 and
4 days of incubation. Fluorescent pseudomonads were
screened under UV (313 nm).
HSL to both rhizospheric (WT plants) and bulk soil,
and re-extracting as indicated above. The detection limit
of this method was estimated to ca. 5 pmol g 1 dry soil.
2.3. Isolation of representative colonies and conservation
Ten lL of an overnight culture of the isolate to assay
in TY was transferred into a well of a microtiter plate
containing 190 lL of TY buffered at pH 6.5 and incubated at 24 C. After 48 h, presence of N-AHSL was assayed with the two biosensors, as described in previous
sections. In the case of N-AHSL degradation tests, 190
lL of liquid TY buffered at pH 6.5 containing 10 lM
of C6-HSL was dispensed into the wells of a sterile
microtiter plate. Ten lL of an overnight culture was
inoculated into each well of the plate, which was then
incubated at 24 C for 24 h. To determine N-AHSL-inactivating activity, 10 lL of the content of each well was
taken. The presence of C6-HSL remaining in the 10 lL
of sample was determined as described above. A control
experiment using non-inoculated degradation medium
was carried out in parallel with the inoculated degradation assays. When appropriate, strains degrading
N-AHSL were assayed for a possible inhibition of
C. violaceum CV026 by a reverse test using 0.25 lM
C6-HSL, as described by Mc Clean et al. [17].
Isolation of bacteria as pure cultures is required to
determine the ability of these bacteria to produce or degrade N-AHSL. At each sampling time, from the isolates obtained from each of the five analyzed pots, 50
colonies were randomly picked from TY plates and 50
colonies from the KBm plates. Per sampling date, a total
of 1000 CFU were therefore isolated (2 analyzed populations, 2 plant genotypes, 5 pots, 50 isolates per pots).
Purified isolates were inoculated into 200 lL TY and
incubated for three days at 24 C, and then stored in
microtitration plates in this medium supplemented with
25% glycerol, at 70 C.
2.4. Detection of N-AHSL
Because N-AHSL are sensitive to alkaline pH [16], all
production and degradation assays used TY media buffered at pH 6.5 with 15 mM KH2PO4/K2HPO4 (to avoid
salt precipitation, CaCl2 was omitted). Detection of NAHSL with Chromobacterium violaceum CV026 [17]
was carried out essentially according to Reimmann
et al. [8]. Detection of N-AHSL with Agrobacterium
tumefaciens NTLR4 [18] on semi-solid media or on
TLC silica plate (Whatman, C18-reverse phase) was carried out essentially according to Shaw et al. [19] and Elasri et al. [20]. Detection limit using the Agrobacterium
biosensor was 0.04 pmol per spot.
2.5. Visualization and quantification of N-AHSL produced by transgenic plants
N-AHSL production by transgenic plants was assessed both in vitro and in situ. In vitro, tobacco plants
were cultivated in 125 mL of sterile MS/2 medium (Sigma,
catalog ref. M11225) in a growth chamber at 24 C under
a photoperiod of 12 h. During 1.5 month, each week, 5
mL of the culture medium was taken, replaced by the
same volume of fresh medium, and extracted twice with
5 mL of ethyl acetate. In situ, 20 g of rhizospheric soil
was extracted twice with 20 mL ethyl acetate. The extracts
were dried over anhydrous sodium sulfate and concentrated by evaporation under a stream of air. The resulting
extracts and 3-oxo-C6-HSL standards were spotted onto
TLC silica plate. The detection of N-AHSL was assessed
with the biosensor A. tumefaciens NTLR4 as indicated
above. Adsorption and natural degradation controls
were also performed by adding known amounts of C6-
2.6. Identification of N-AHSL-producing and N-AHSLdegrading bacteria
2.7. PLFA analyses
The samples analyzed (30 g) by PLFA consisted in
the root system of both WT and transgenic tobacco
plants, plus some residues of rhizosphere soil adhering
to the roots, harvested at 15 weeks. Analyses were performed in triplicate by Microbial Insights (http://
www.microbe.com/). Lipids were recovered using a
modification of the method of Bligh and Dyer, according to White et al. [21]. Extractions were performed
using one-phase chloroform–methanol buffer extraction.
Recovered lipids were dissolved in chloroform and fractionated on disposable silicic acid columns into neutral,
glyco, and polar lipid fractions. The polar lipid fraction
was trans-esterified with mild alkali to recover the PLFA
as methyl esters, in hexane. PLFA were analyzed by gas
chromatography with peak confirmation performed by
electron impact mass spectrometry (GC/MS). PLFA
nomenclature follows the pattern A:BxC. The ‘‘A’’ position identifies the total number of carbon atoms in the
fatty acid. Position B is the number of double bonds
from the aliphatic (x) end of the molecule. Position
‘‘C’’ designates the carbon atom from the aliphatic
end before the double bond. This is followed by a ‘‘c’’
for cis or a ‘‘t’’ for trans configuration. The prefixes
‘‘i’’ and ‘‘a’’ stand for iso and anteiso branching. Midchain branching is noted by ‘‘me’’, and cyclopropyl-fatty
acids are designated as ‘‘cy’’ [22].
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2.8. DGGE analysis of bacterial regions encoding rRNA
The structure of the bacterial population associated
with the root system (ca. 10 g) of both WT and transgenic plants producing N-AHSL was analyzed by
DGGE of PCR-amplified DNA regions encoding
rRNA (rrs gene). DGGE analyses were sub-contracted
to Microbial Insights (http://www.microbe.com/). Nucleic acid extraction was performed using a bead-beating method [23]. The DNA was purified by a glassmilk DNA purification protocol using a Gene CleanTM
kit as described by the manufacturer. PCR amplification
of 16S rRNA gene fragments was performed as
described by Muyzer et al. [24] with modifications.
Thermocycling consisted of 35 cycles of 92 C for 45 s,
55 C for 30 s, and 68 C for 45 s. Using 1.25 units of
Expand High Fidelity polymerase and 10 pmol each primer (forward primer contained a 40 bp GC-clamp) in a
total volume of 25 lL, thermocycling was performed
using a ‘‘RobocyclerTM’’ PCR block. The primers targeted eubacterial 16S rDNA regions corresponding to
Escherichia coli positions 341–534. DGGE employed a
D-Code 16/16 cm gel system maintained at a constant
temperature of 60 C in 6 L of 0.5· TAE buffer (20
mM Tris actate, 0.5 mM EDTA, pH 8.0). Denaturing
gradients were formed between 30% and 65% denaturant (with 100% denaturant defined as 7 M urea, 40%
v/v formamide). Gel images were captured using an Alpha ImagerTM system. Purified DNA was sequenced
with an ABI-Prism automatic sequencer model 377 with
dye terminators. Sequence identifications were performed using the BLASTN facility of the National Center for Biotechnology Information (http://ncbi.nlm.nih.
gov/Blast) and the ‘‘Sequence Match’’ facility of the
Ribosomal Database Project (http://www.cme.msu.edu/
RDP/analyses.html). Gel images were standardized and
examined visually. Band positions (Rf) in each lane
were converted to 0 (absence) and 1 (presence) values
and assembled in a matrix. Profile similarity was calculated with the Dice algorithm using the DistAFLP
software (http://pbil.univ-lyon1.fr/ADE-4/microb). A
dendrogram was constructed by using the unweighted
pair group method with mathematical average (UPGMA) using the Phylip package and the Treeview software, (http://taxonomy.zoology.gla.ac.uk/rod/rod.html).
StudentÕs t test (P > 0.05) on the average enumeration
values obtained for each of the microcosms.
3. Results and discussion
3.1. N-AHSL production by the transgenic tobacco plant
line
As a prerequisite to this study, production of C6HSL and 3-oxo-C6-HSL by plants was verified in vitro
and assessed in soil. In soil, presence of 3-oxo-C6-HSL
was determined in two rhizosphere zones, ca. 3.5 months
after seed germination. Taking into account the degradation inherent to the soil microbial activity, the adsorption on soil particles and the extraction yield, the
assayed N-AHSL (3-oxo-C6-HSL) was present at ca. 5
pmol g 1 dry soil in the ‘‘outer’’ rhizosphere, the rhizosphere zone located far from the plant, that contains
mostly soil and very little roots. The molecule was detected at 2 nmol g 1 dry soil in the rhizosphere soil that
contains a dense plant root system. No N-AHSL production was detected at the root system of WT tobacco.
Furthermore, the transgenic tobacco plants and the WT
line showed identical morphology and growth parameters in the microcosms. The measured concentrations
cannot be compared with published data because most
of these data were obtained in vitro, while a non-sterile
rhizosphere constitutes an ‘‘open environment’’. However, values comparable to those indicated above have
been reported in soil [25] and in a tomato rhizosphere
(40 nM, i.e. ca. 0.5 nmol g 1 dry soil [26]).
In vitro, in MS/2 medium, N-AHSL production was
linear over a 6-week period and reached ca. 4.5 pmol of
equivalent 3-oxo-C6-HSL mL 1, i.e. ca. 8 nmol of
equivalent 3-oxo-C6-HSL g 1 dry root, after 6 weeks
of plant growth. In addition, under those conditions,
inoculation of the Chromobacterium sensor CV026 at
the surface of the roots induced violacein production
(not shown). Similarly, presence of N-AHSL could be
demonstrated in the root system of the yenI plants that
were grown in the experimental soil and harvested
(Fig. 1). This is consistent with previous data indicating
that these transgenic plants cross-talk with both Erwinia
and Pseudomonas isolates [4,10]. All these data demonstrate that N-AHSL were produced at biologically active concentrations in the plant environment.
2.9. Statistical analysis
3.2. Analysis of bacterial groups inhabiting the bulk soil
For each condition, at each time point, five microcosms (each with three plants) were individually analyzed. Enumerations were performed in triplicate for
each microcosm from the same root and soil sample.
Potential outliers were detected using GrubbsÕ ESD test
(P > 0.05). From one condition to another (time or
plant line), comparison of means was performed using
The bacterial populations examined in this study
were the total culturable microbial community and some
groups were chosen because several of their members exhibit, or respond to QS regulation (e.g. fluorescent Pseudomonas). In addition, the cell densities of bacterial
communities implicated in QS either as producing N-
C. dÕAngelo-Picard et al. / FEMS Microbiology Ecology 51 (2004) 19–29
23
after 7, 11 and 15 weeks. Results are shown in Fig. 2.
They revealed that the density of total cultivable bacteria remained identical or near-identical from 7 to 15
weeks, in the rhizosphere of WT plants (Fig. 2(a)). Similar results were found for the densities of thermotolerant bacteria and fluorescent peudomonads in the
rhizosphere of WT plants, and for the densities of the
same groups in the rhizosphere of plants producing
N-AHSL (Fig. 2(b)). Comparison of the bacterial populations in the rhizosphere of the two plant lines with
8
7
3.3. Enumerating bacterial populations from WT and NAHSL-producing tobacco rhizosphere
Further enumerations of bacterial populations were
performed from the rhizosphere of both WT and transgenic tobacco producing C6-HSL and 3-oxo-C6-HSL,
a
*
a b ab
6
5
a b c
abc
c
a
b
b
b
4
3
2
1
0
total viable
bacteria
(a)
B
9
8
log (UFC/g soil)
AHSL signals or degrading the N-AHSL C6-HSL were
also monitored. Importantly, the designation ‘‘degradation’’ or ‘‘degrader’’ (as well as that of ‘‘quencher’’) does
not imply that the relevant isolates assimilated C6-HSL
as a carbon and energy source.
Bacterial populations were examined in the original
soil mixture (time zero) just prior to seed germination.
Results are shown in Fig. 2. The total cultivable, bacterial population reached 6 · 106 CFU g 1 of soil mixture,
while fluorescent pseudomonads occurred at ca. 8 · 104
CFU g 1, sporulated bacteria at ca. 2 · 105 CFU g 1
and thermotolerant at ca. 3 · 105 CFU g 1 soil. At time
zero, 250 isolates taken out of the total microbial
community and 250 isolates taken out of fluorescent
pseudomonads were subjected to N-AHSL production
and C6-HSL degradation assays. Amongst total viable
and cultivable bacteria, those producing N-AHSL represented 8% of the assayed population, while ca. 4%
quenched the C6-HSL signal. Amongst fluorescent pseudomonads, the corresponding figures for these communities were 30% and below the detection limit (i.e. 0.4%),
respectively, as no C6-HSL-degrading strains were
detected amongst the 250 Pseudomonas soil isolates.
b c
bc
9
log (UFC/g soil)
Fig. 1. Evidence for the release of quorum-sensing signal molecules by
the roots of the tobacco plants expressing the yenI gene. Transgenic
and WT plants expressing yenI were cultivated in the non-sterile
experimental soil for 11 weeks. At this time plants were harvested, soil
particles removed by vigourous shaking in the air, and the roots placed
onto AB medium containing X-gal (40 lg mL 1) for 12 h. (a) Plant
roots were laid onto Agrobacterium biosensor NTLR4, that detects the
production of N-AHSL. (b) Control performed without biosensor,
allowing the visualization of the putative b-galactosidase activity of the
plant-associated microorganisms. A large N-AHSL production zone is
seen around roots of the yenI line, while very limited production zones
are detected around the roots of the WT line (a). Both observations
cannot be attributed to the production of beta-galactosidase by the
plant roots and the bacteria (b).
7
B
heat resistant,
spore-forming
bacteria
thermotolerant
Fluorescents
Pseudomonads
B
A
6
AB
5
*
A
A
A A
CC
B B B
A
4
3
2
1
0
(b)
total viable
bacteria
heat resistant,
spore-forming
bacteria
thermotolerant
Fluorescents
Pseudomonads
Fig. 2. Dynamics of various bacterial populations obtained from soil
samples, and from the rhizosphere of WT and genetically modified
tobacco producing N-AHSL. Microbial populations and communities
(total cultivable, sporulated, thermotolerant, and fluorescent pseudomonads) were enumerated from bulk soil samples (white bars) and
from the rhizosphere of tobacco, at 7 weeks (light grey bars), 11 weeks
(dark grey bars) and 15 weeks (black bars). Each bar represents the
average log number of bacteria per gram of soil calculated from
enumeration performed in triplicate, from five independent microcosms per condition. Within studied groups, the values marked with
the same letter did not differ significantly upon time, one from the
other, based on StudentÕs t test (P > 0.05). Lower case letters,
comparison of values for bacteria isolated from WT tobacco; upper
case letter, for bacteria isolated from transgenic tobacco producing NAHSL. The symbol * identifies the values that differed from one plant
environment (WT) to the other (N-AHSL-producing tobacco), based
on StudentÕs t test (P > 0.05). (a) WT tobacco; (b) transgenic tobacco
producing N-AHSL.
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C. dÕAngelo-Picard et al. / FEMS Microbiology Ecology 51 (2004) 19–29
those detected in the original soil sample revealed that
the density of total viable bacteria and that of fluorescent pseudomonads increased during seed germination
and early developmental steps of the plant, while the
density of heat-resistant, sporulated bacteria remained
stable or slowly decreased. This phenomenon is known
as the rhizosphere effect, itself related to rhizodeposition (i.e. the release of bacterial growth substrates into
the rhizosphere) that results from the photosynthetic
and metabolic activities of the plant [27].
The values obtained at each time point for each
microbial group were compared as a function of the
plant line. Whatever the time point (7–15 weeks), the
average values of all assayed populations did not significantly differ (P > 0.05) from the rhizosphere of one
plant line to that of the other, excepted for sporulated
bacteria enumerated at 11 weeks. No clear biological
explanation can be proposed to account for this
observation.
3.4. Structure of cultivable and non-cultivable microbial
populations isolated from WT and N-AHSL-producing
tobacco rhizospheres
Microbial populations and communities were analyzed at 15 weeks using PLFA and DGGE analyses on
samples originating from the rhizosphere of both WT
and N-AHSL-producing plants.
The PLFA profiles were carried out in triplicate.
Average results (and standard deviation) are given in
Table 1. Whatever the plant line, the samples contained
ca. 75 nmol of PLFA g 1, corresponding to ca.
1.5 · 109 cells g 1. This value is about 10 times higher
than that obtained above when enumerating cultivable
bacteria. This ratio (cultivable/total = 0.1) appears consistent with results reported by others in comparable
systems [28]. In both types of rhizosphere, the values
of the bacterial markers for cell turnover rate (Cy/
x7c) showed that the bacterial community was suffering a standard degree of starvation for a soil community. The toxic stress markers trans/cis 16:1x7 and
18:1x7 were low, indicating that the bacterial communities were not affected by the presence of compounds
with toxic or antibiotic activities. Overall, microbial
biomass measurements, starvation and stress indexes
were not significantly different (P > 0.05) from one
rhizosphere to the other (WT vs. N-AHSL-producing
plants).
Aside from the ubiquitous 16:0 fatty acid, the most
abundant species in the samples was the mono-unsaturated acid 18:1w7c (data not shown), common in
Gram-negative bacteria [29] that are frequent inhabitants of the rhizosphere. The most abundant components
of the samples were eukaryotes, possibly from fungal
and plant origin as these samples consisted in roots
and tightly associated soil particles. Amongst bacteria,
aerobic and aero-anaerobic gram-negative ones were
Table 1
PLFA analysis of the microbial community colonizing the root environment of WT and transgenic plants
Properties of the microbial community
Wild-type plants
N-AHSL-producing plants
(a) Biomassa
Total amount PLFA (nmol g 1)
Cell equiv. value (cells g 1)
Bacterial PLFA (nmol g 1)
Eukaryote PLFA (nmol g 1)
Ratio bacterial PLFA/eukaryote PLFA
83.3 (26.5)b
1.67 · 109 (5.30 · 108)
41.8 (9.8)
41.5 (16.8)
1.07 (0.24)
68 (16.9)
1.36 · 109 (3.39 · 108)
39.2 (7.9)
28.8 (9.2)
1.40 (0.22)
(b) Metabolic activity (Cy/x7c)
Marker A (ratio cy17:0/16:1w7c)
Marker B (ratio cy19:0/18:1w7c)
A + B index
0.08 (0.02)
0.14 (0.01)
0.21b (0.02)
0.11 (0.01)
0.15 (0.01)
0.26 (0.02)
0.04 (0.01)
0.03 (0.00)
4.43 (1.39)
18.88 (5.66)
1.25 (0.50)
1.16 (0.39)
48.65 (5.43)
25.64d (2.49)
6.17 (0.8)
25.15 (3.67)
1.89 (0.40)
1.68 (0.38)
41.88 (3.66)
23.25d (1.63)
(c) Environmental stress
Gram-negative population (16:1w7t/16:1w7c)
c
(d) Community structure (% of total PLFA)
Gram-positive and anaerobic gram-negative (terminally branched saturated)
Gram-negative (monoenoic)
Anaerobic metal reducers (branched monoenoic)
Sulfate-reducing bacteria and Actinomycetes (mid-chain branched saturated)
Eukaryotes (fungi, protozoa, algae, etc.) (polyenoics)
Miscellaneous (saturated)
a
All biomass values are given per gram of dry root and soil mixture. Statistical analysis indicated that none of the values presented in the middle
column (WT plants) differed (P > 0.05) from the values presented in the right column (N-AHSL-producing plants).
b
Average values obtained from three independent repeats are presented, with standard deviations in between parentheses.
c
Total is different from 0.22 due to rounding.
d
Total is different from 100% due to rounding.
C. dÕAngelo-Picard et al. / FEMS Microbiology Ecology 51 (2004) 19–29
about four times more abundant than gram-positive and
anaerobic gram-negative bacteria. Most importantly,
none of the proportions presented in Table 1 differed
statistically (P > 0.05) from one rhizosphere to the
other. Taken together with the above data, the results
suggest that the production of QS signal molecules by
the plant affected neither the structure of the eukaryotic
and prokaryotic populations, nor part of the microbial
physiology related to starvation and toxic stress, at least
at the time the analyses were performed.
Bacterial populations associated with the root system
of both WT and N-AHSL-producing plants were also
analyzed using DGGE of PCR-amplified DNA regions
encoding rRNA. Results, obtained on three independent
experiments, are shown in Fig. 3. The DGGE profiles
appeared complex, with a minimum of at least 30 bands
per lane. This apparent complexity was previously reported by other authors who studied soil-borne or
root-associated bacterial populations [30,31]. Several
bands were excised and sequenced. Sequence analysis allowed the identification of rRNA genes having homology to both a- and b-proteobacteria, which are both
known to be plant surface colonizers. Several bacteria
belonging to the Bacteroidetes group that encompasses
the super families Bacteroides, Flavobacteria, and
Sphingobacteria were identified. These are typically bacteria that may have escaped the enumeration in the culture-based approach used in this study as some of these
25
are strictly anaerobic organisms. No conclusion can be
drawn with respect to the relative abundance of these
groups one to the other as the intensity of the bands generated by the PCR amplification is not directly related to
the relative abundance of the targeted group [32]. Interestingly, sequences characteristics for the chloroplastic
genome of Nicotiana plant species have been identified
in this analysis, a feature, which constitutes an impromptu positive control validating the technique.
Direct examination of the DGGE profiles did not allow the identification of any amplified fragment that
would be specific for WT plants or for transgenic plants
expressing yenI. However, differences were observed
from one sample to another. To investigate whether
these variations were due to sampling per se or related
to the plant genotype, the six profiles were converted
into a 0/1 matrix and an UPGMA tree representing
the genetic similarity of the microbial community was
built. The tree, shown in Fig. 3, reveals that a sample
originating from WT plants may be as distant from samples originating from other WT plants than from samples originating from plants producing QS molecules
(T-plants). Similarly, the microbial community originating from a T-plant may be more related to that originating from a WT plant than from another T-plant.
Overall, the tree indicates that the various patterns observed are not related to the plant phenotype but rather
to experimental variability. In other words, the
Fig. 3. DGGE profiles of PCR-amplified DNA regions encoding 16S rRNA obtained from the root-associated microbial community. DGGE
patterns were obtained from three samples obtained independently from wild-type (WT1–3) plants or from N-AHSL-producing plants (T1 to T3).
Dendrograms were constructed from the DGGE community fingerprints as indicated in Section 2. Determination of pattern similarity was based on
cluster analysis with the DiceÕs algorithm. Labelled bands (marked A to G) were excised and sequenced; results are provided in the figure along with
the percentage of homology to the closest taxonomic group.
26
C. dÕAngelo-Picard et al. / FEMS Microbiology Ecology 51 (2004) 19–29
production of the QS signal molecules C6-HSL and 3oxo-C6-HSL does not appear to alter the composition
of the soil microbial community. This conclusion is consistant with that derived from the PLFA analysis.
tobacco plants. The values obtained at each time point
for each microbial group were compared as a function
of the plant line. Whatever the time point (7–15 weeks),
the average occurrence of N-AHSL-producing strains
amongst the total cultivable microbial community did
not significantly differ from the rhizosphere of one plant
line to that of the other plant line (P > 0.05). Amongst
the pseudomonads, the average occurrence of NAHSL-producing strains differed from one plant line
to the other only at 7 weeks.
The proportion of N-AHSL producers in the rhizosphere of WT tobacco is not significantly different from
that of N-AHSL producers inhabiting the bulk soil.
Interestingly, the frequency of N-AHSL-producing
pseudomonads averages ca. 15% in the rhizosphere of
WT tobacco. This ratio is not significantly different
from, or lower than that observed for the same population isolated from bulk soil and contrasts a previous result [20]. The discrepancy is probably due to the fact that
the above results were obtained using a collection of
Pseudomonas strains and soils of different characteristics
and origins.
The high proportion of bacteria exhibiting N-AHSLbased QS regulation reported in this work is in
3.5. Enumerating N-AHSL-producing bacteria from WT
and N-AHSL-producing tobacco rhizosphere
N-AHSL-producing bacteria were obtained from the
rhizosphere of the different plant lines and their respective occurrence was compared. Results are given in Figs.
4(a) and (b). The occurrence of strains producing NAHSL, isolated from the cultivable microbial community colonizing WT tobacco, remained constant over
the duration of the experiment, from 7 to 15 weeks
(P > 0.05), while that of the strains producing N-AHSL
obtained from the rhizosphere of N-AHSL-producing
tobacco plants appeared to vary more drastically.
Enumerations performed on Pseudomonas strains isolated from the rhizosphere of WT tobacco revealed that
the occurrence of N-AHSL-producing pseudomonads
remained constant over the duration of the experiment,
from 7 to 15 weeks (P > 0.05), as did that of the isolates
obtained from the rhizosphere of N-AHSL-producing
(b)
50
40
B
30
20
B
b
ab
10
ab
a
A
0
0
A
5
10
% HSL producing bacteria
% HSL producing bacteria
(a)
50
40
a
A
30
ab
20
ab
B
10
B
b
B
0
15
0
5
time (weeks)
10
15
time (weeks)
%HSL degrading bacteria
(c)
50
40
30
20
A
10
a
A
0
0
5
a
A
a
A
a
10
15
time (weeks)
Fig. 4. Dynamics of N-AHSL-producing and -degrading bacteria in soil. The ratio of bacteria-producing or -degrading N-AHSL was determined as
indicated in Section 2, from soil samples (open square marks) and from the rhizosphere of tobacco, at 7 weeks, 11 weeks and 15 weeks (circles and
triangles). Each ratio was calculated from the average number of bacteria per gram of soil producing or degrading N-AHSL, obtained from
enumeration performed on 250 isolates, i.e. 50 isolates obtained from five independent microcosms per condition. Grey circles represent the values
obtained from WT tobacco and black triangles, the values obtained from transgenic tobacco producing N-AHSL. The values marked with the same
letter did not differ significantly one from the other, based on StudentÕs t test (P > 0.05). Lower case letters, comparison of values for bacteria isolated
from WT tobacco (grey circles); upper case letter, comparison of values for bacteria isolated from transgenic tobacco producing N-AHSL (black
triangles). (a) N-AHSL-producing bacteria within total, cultivable microbial community; (b) N-AHSL-producing bacteria within fluorescent
pseudomonads; and (c) N-AHSL-degrading bacteria within total, cultivable microbial community.
C. dÕAngelo-Picard et al. / FEMS Microbiology Ecology 51 (2004) 19–29
agreement with the values obtained by other researchers
(12% [26]) for plant-associated bacteria. However, lower
values for the occurrence of N-AHSL-producing microorganisms in the rhizosphere have also been reported
(8% [33]; ca. 10% [34]). This is possibly related to the
use, by these authors, of only one N-AHSL biosensor
to identify the N-AHSL-producing strains and to the
fact that all available biosensors exhibit some specificity
towards the N-AHSL molecules [17,18].
3.6. Enumerating C6-HSL-degrading bacteria from WT
and N-AHSL-producing tobacco rhizosphere
Bacteria degrading the C6-HSL signal were obtained
from the rhizosphere of the different plant lines and their
respective occurrence was compared (Fig. 4(c)). The
occurrence of C6-HSL-degrading strains isolated from
the rhizosphere of both WT and N-AHSL-producing tobacco plants did not vary with time (from time zero to
15 weeks – P > 0.05) and averaged 5–10% amongst cultivable microorganisms. The values obtained at each
time point were compared as a function of the plant line.
Regardless of the time point (7–15 weeks), the average
occurrence of N-AHSL-degrading strains did not significantly differ from the rhizosphere of one plant line to
that of the other plant line (P > 0.05).
Enumerations performed on fluorescent pseudomonads yielded only 2 C6-HSL-interfering isolates. Both
were obtained at 7 weeks, one from the rhizosphere of
WT plants and the other from the rhizosphere of tobacco
plants producing N-AHSL. The very low frequency of
pseudomonads degrading C6-HSL does not allow statistical comparison between the two plant lines.
Overall, the proportion of cultivable bacteria interfering with the N-AHSL signal reported in this work is in
agreement with the values obtained by others in other
soils or plant environments (ca. 2% [8]; 5–7% [7,35]). Similarly, the occurrence of Pseudomonas strains degrading
N-AHSL signals has already been reported, whether
these were soil-borne or plant-associated bacteria [9,36].
3.7. Relationships of N-AHSL-producing and C6-HSLdegrading communities
This study provides a comparison of microbial communities involved in QS regulation, either as producers
or as degraders, in soil and in a plant rhizosphere. As
stated above, out of 250 isolates representing the total
microbial community isolated from bulk soil (time zero),
20 (8%; Fig. 4(a)) produced at least one N-AHSL QS
signal molecule. Interestingly, none of these 20 isolates
degraded C6-HSL. Similarly, out of 250 fluorescent
pseudomonads isolated from bulk soil, 75 (30%; Fig.
4(b)) produced at least one N-AHSL, but none of these
degraded the C6-HSL signal molecule. A similar analysis was performed at 7 weeks, 11 weeks, and 15 weeks,
27
on isolates obtained from the rhizosphere of both plant
lines. None of the producing isolates were found to degrade C6-HSL and vice versa.
Results described above yield novel ecological data:
they indicate that, at least in the investigated plant environment, N-AHSL-producing and N-AHSL-interfering
communities do not encompass the same sub-populations. As a consequence, the N-AHSL signal may indeed
constitute a valuable target for competition in the rhizosphere. From this, however, a question arises: does the
natural production of these molecules by microorganisms with QS regulation suffice to explain the relatively
high frequency of interfering strains in the soil and the
rhizosphere? In terms of selective pressure, the benefit
most probably does not rely upon a trophic advantage
held by the degrading strains, because N-AHSL concentration is very low in the rhizosphere and also because
several of the quenchers isolated in a different work did
not assimilate N-AHSL or N-AHSL breakdown products (Uroz, unpublished data). Rather, this benefit might
rely upon an increased competitiveness of the degraders
vs. other rhizosphere microorganisms, via a ‘‘natural
quenching effect’’. However, it cannot be excluded that
N-AHSL degradation may be fortuitous amongst bacteria, and not primarily directed towards the disruption of
QS signal molecules, the relevant enzymes perhaps catalyzing alternative reactions [37]. In addition, N-AHSL
catabolism may be part of a more complex degradative
network, as reported earlier [38].
3.8. Concluding remarks
Though present at low concentrations in the environment, N-AHSL are potent signal molecules common
among plant-associated bacteria [18], that diffuse in
the rhizosphere and affect key functions of defined bacterial groups [4,12,26]. This study demonstrates that the
release of C6-HSL and 3-oxo-C6-HSL in the rhizosphere of a tobacco line has no or very limited consequences on the root-associated microbial community,
including cultivable bacteria whether the latter belonged
to reporter groups (pseudomonads, thermotolerant,
sporulated bacteria) or communities involved in QS regulation. Based on the results described above, it appears
that biocontrol strategies relying upon the saturation of
a plant environment with N-AHSL molecules may induce no, or very limited disturbance of the plant-associated microbial community. This conclusion should now
be challenged in additional plant and soil systems.
Acknowledgments
The authors thank the EU for the support provided
to this work via the Eco-safe program granted to R.F.
and Y.D. This work was also supported by the
28
C. dÕAngelo-Picard et al. / FEMS Microbiology Ecology 51 (2004) 19–29
programs ‘‘OGM’’ from INRA and ‘‘Impact des nouvelles technologies’’ from CNRS to Y.D., by BRG to
D.F., and by a BBSRC Sir David Phillips fellowship
awarded to R.F. S.U. is a recipient of a fellowship from
Ministère de la Recherche et des Nouvelles Technologies. The authors thank all the members of the Eco-safe
EU consortium for their helpful comments on this work.
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